The Bay of Bengal (BoB) has long stood as a biogeochemical enigma, with
subsurface waters containing extremely low, but persistent, concentrations
of oxygen in the nanomolar range which – for some, yet unconstrained, reason –
are prevented from becoming anoxic. One reason for this may be the low
productivity of the BoB waters due to nutrient limitation and the resulting
lack of respiration of organic material at intermediate waters. Thus, the
parameters determining primary production are key in understanding what
prevents the BoB from developing anoxia. Primary productivity in the sunlit
surface layers of tropical oceans is mostly limited by the supply of
reactive nitrogen through upwelling, riverine flux, atmospheric deposition,
and biological dinitrogen (N2) fixation. In the BoB, a stable
stratification limits nutrient supply via upwelling in the open waters, and
riverine or atmospheric fluxes have been shown to support only less than one-quarter of the nitrogen for primary production. This leaves a large
uncertainty for most of the BoB's nitrogen input, suggesting a potential
role of N2 fixation in those waters.

Here, we present a survey of N2 fixation and carbon fixation in the BoB
during the winter monsoon season. We detected a community of N2 fixers
comparable to other oxygen minimum zone (OMZ) regions, with only a few
cyanobacterial clades and a broad diversity of non-phototrophic N2
fixers present throughout the water column (samples collected between 10
and 560 m water depth). While similar communities of N2 fixers were
shown to actively fix N2 in other OMZs, N2 fixation rates were
below the detection limit in our samples covering the water column between
the deep chlorophyll maximum and the OMZ. Consistent with this, no N2
fixation signal was visible in δ15N signatures. We suggest that
the absence of N2 fixation may be a consequence of a micronutrient
limitation or of an O2 sensitivity of the OMZ diazotrophs in the BoB.
Exploring how the onset of N2 fixation by cyanobacteria compared to
non-phototrophic N2 fixers would impact on OMZ O2 concentrations,
a simple model exercise was carried out. We observed that both photic-zone-based and OMZ-based N2 fixation are very sensitive to even
minimal changes in water column stratification, with stronger mixing
increasing organic matter production and export, which can exhaust
remaining O2 traces in the BoB.

Primary production in large areas of the surface ocean is limited by the
availability of fixed nitrogen (Moore et al., 2013). This deficiency in
nitrogen creates a niche for dinitrogen (N2) fixation, an energy-costly
process carried out only by certain prokaryotes, also referred to as
diazotrophs, which are phylogenetically highly diverse. N2 fixation in
the ocean has been described quantitatively as most important in the
oligotrophic surface waters of the subtropical gyres (Sohm et al.,
2011; Luo et al., 2012; Wang et al., 2019) where cyanobacterial N2 fixers
dominate. Over the last decade, the development of novel molecular tools
revealed that non-cyanobacterial N2 fixers are widely distributed
throughout ocean waters (Farnelid et al., 2011,
2013; Fernandez et al., 2011; Luo et al., 2012; Riemann et al., 2010; Zehr et
al., 1998) and sediments (Fulweiler et al., 2007; Andersson et al.,
2014; Bertics et al., 2013; Gier et al., 2017, 2016). Their
quantitative importance for global N2 fixation, however, is not yet
clear. In oxygen minimum zones (OMZs) of the eastern tropical North and South Pacific Ocean,
hypoxic basins in the San Pedro Ocean Time-series and the Santa Monica Bay Observatory in the Southern California Bight, and the Arabian Sea, those N2
fixers form a unique community consisting of different clades of
proteobacteria, clostridia, spirochaetes, chlorobia, and methanogenic
archaea (Christiansen and Loescher, 2019; Dekaezemacker et al.,
2013; Fernandez et al., 2011; Gaby et al., 2018; Gier et al., 2017; Goebel et
al., 2010; Halm et al., 2012; Hamersley et al., 2011; Jayakumar et al.,
2012, 2017; Löscher et al., 2014). In contrast,
cyanobacterial N2 fixers and diatom–diazotroph associations (DDAs),
which are commonly considered the most important N2 fixers in the
surface ocean, were either absent or were detected only in low abundances in
OMZs (Turk-Kubo et al., 2014; White et al., 2013; Jayakumar et al., 2012).
Both the presence of diazotrophs clustering with proteobacteria,
clostridia, spirochaetes, chlorobia, and methanogenic archaea and the
underrepresentation of cyanobacterial and DDA N2 fixers could thus be
considered characteristic for OMZ environments.

Nutrient stoichiometry and model predictions (Deutsch
et al., 2007) suggest that oxygen-deficient waters are a potentially
important niche for N2 fixation. Based on this suggestion, several
studies have focused on N2 fixation in the large and persistent OMZ of
the eastern tropical South Pacific. In this region N2 fixation rates
vary, however, with maximum rates of 840 µmolNm-2d-1
detected in nitrogenous sulfidic waters off the coast of Peru
(Löscher et al., 2014) and 117 µmolNm-2d-1 in the oxygen-depleted zone (Bonnet et
al., 2013). Low N2 fixation rates close to the detection limit were
reported from the same area (Chang et al., 2019); another set of
N2 fixation rates estimated from sediment trap analyses were in the
range of 0–23 µmolNm-2d-1 (Knapp et al.,
2016). Taken together, these rates suggest either a strong temporal
variation or spatial patchiness. A similar variation in N2 fixation
rates was described for the eastern tropical North Pacific, ranging from
close to the detection limit in the OMZ (Jayakumar et al.,
2017) up to 795 µmolNm-2d-1 in episodic diazotroph
blooms (White et al., 2013). This apparent temporal or
spatial variation in N2 fixation rates may originate from unresolved
environmental controls on N2 fixation and makes it difficult to
quantify N2 fixation in OMZ waters.

The Bay of Bengal (BoB) is a region with a strong seasonality driven by the
Asian monsoon system. Massive rainfall and river discharge with maximum
freshwater inputs in September (e.g., Mahadevan, 2016) cause a
considerable lowering of surface water salinity during and after the
monsoons throughout the whole basin (Subramanian, 1993). This,
together with increased surface water temperatures, leads to intensive and
persistent stratification of the water column (Kumar et al.,
2004), restricting nutrient fluxes to the surface from below and promoting a
strong OMZ (McCreary et al., 2013; Paulmier and Ruiz-Pinto, 2009; Sarma,
2002) with minimum oxygen (O2) concentrations in the lower nanomolar
range (Bristow et al., 2017).

The potential importance of N2 fixation in the BoB can be derived from
a simple N budget estimate with an overall N loss of 7.9±0.6Tg N yr−1 and N sources other than N2 fixation of 3.15±2.25Tg N yr−1 (Table 1 – data from Naqvi, 2008; Naqvi et al., 2010; Bristow
et al., 2017; Singh et al., 2012; Krishna et al., 2016; Srinivas and Sarin,
2013; Suntharalingam et al., 2019). This implies a deficit of 4.7±2.4Tg N yr−1 within the given range of uncertainty, indicating the
potential importance of N2 fixation, assuming a coupling of nitrogen
loss and N2 fixation as proposed by Deutsch et
al. (2007). Naqvi et al. (2010) proposed N2 fixation to contribute 1 Tg N yr−1 in the BoB, while Srinivas and Sarin (2013) interpolated a
contribution of 0.6–4 Tg N yr−1 from phosphate availability.
Measurements of N2 fixation rates from the BoB are not available,
and isotope analysis of sediment trap samples indeed suggests that the BoB is a
site of active N2 fixation. The composition of the organic
material produced in BoB surface waters is characterized by a high portion
of biogenic opal (20 %) and a low δ15N nitrate signal (3.2 ‰–5 ‰; Gaye-Haake et al., 2005). This
points towards a production of a considerable part of organic matter
produced by diatoms in symbiotic association with or in close proximity to
diazotrophs (Subramaniam et al., 2008). Only few
studies report the presence of diazotrophs including Trichodesmium in the BoB (Wu et
al., 2019; Shetye et al., 2013; Sahu et al., 2017; Jyothibabu et al.,
2006; Mulholland and Capone, 2009), with only one of them using a functional
gene approach.

Table 1Nitrogen fluxes in the BoB (Tg N yr−1). N loss fluxes are
given in black, and N sources are given in gray. DIN: dissolved inorganic
nitrogen. PON: particulate organic nitrogen. IO: Indian Ocean. N2
loss by denitrification was excluded by
Bristow et al. (2017). Naqvi et al. (2010) reported possible N loss to the atmosphere in the form of N2O.

To investigate the diazotrophic community and to quantify N2 and carbon
fixation in the BoB OMZ, we used a combination of gene sequencing and
quantification, rate measurements, isotope tracing, and box modeling.

2.1 Geochemical sampling

Samples were collected from the top 500 m of the water column during the
SK-308 cruise with the ORV Sagar Kanya to the BoB during the winter monsoon between 24 January and 3 February 2014. Seawater samples were collected using 5 and
30 L Niskin bottles on a CTD rosette equipped with a Sea-Bird SBE 43 oxygen
sensor and a Wet Labs ECO-AFL/FL chlorophyll sensor as previously described
in Bristow et al. (2017). To resolve
oxygen dynamics below the Sea-Bird sensor's detection limit, a STOX
(switchable trace oxygen) amperometric oxygen sensor was used
(Revsbech et al., 2009), which had a detection limit of 7–12 nmol L−1 during this sampling campaign
(Bristow et al., 2017). Nutrients,
including nitrate, nitrite, and phosphate, were determined according to
Grasshoff et al. (1999).

2.2N2–C fixation rate measurements

Seawater was collected from depth between 60 and 280 m water depth. Water
was taken from Niskin bottles and filled into 2.4 L glass bottles or 2.8 L
polycarbonate bottles for (near-)anoxic and all other (oxic) waters,
respectively. Bottles were capped with black rubber stoppers (anoxic waters)
or Teflon-coated butyl rubber septa (oxic waters). Incubations were
performed with the method developed by Mohr et al. (2010), as described in
Grosskopf et al. (2012). Batches of 15N2-gas-enriched (Cambridge
Isotopes, USA) water were prepared with degassed water from two to three of the six sampling depths. Each incubation bottle was supplemented
with 50 mL of the 15N2-enriched seawater. Discrete samples for the
measurement of the 15N2 concentration were taken from each
incubation bottle and were measured by membrane-inlet mass spectrometry
(MIMS). Final 15N2 enrichments were on average 1.65 at. %
15N. For carbon fixation measurements, NaH13CO3 was dissolved
in sterile Milli-Q water (1 g∕117 mL), and 5 mL was added to each incubation
(∼8 at. % final, based on total DIC of 2.2 mM). Bottles
with water from the upper two depths were kept in surface seawater-cooled
on-deck incubators. Bottles from the lower depths were incubated at
13–15 ∘C in the dark. Incubations were stopped after approximately
24 h (samples with less than 20h incubation time were excluded from our
analysis). Volumes between 2.1 and 2.7 L of seawater were filtered onto
pre-combusted (450 ∘C; 4–6 h) 25 mm diameter GF/F filters
(Whatman, GE Healthcare, Chalfont St Giles, UK) under a gentle vacuum (200 mbar). Filters were either frozen at −20∘C and oven dried prior
to processing or oven dried (50 ∘C) directly for 24 h and stored
dry until analysis. Untreated seawater was filtered and prepared as
described above to obtain background natural abundance values. For elemental
and isotopic analysis, GF/F filters were acidified over fuming HCl overnight
in a desiccator to remove inorganic C. Filters were then oven dried for 2 h at 50 ∘C and pelletized in tin cups. Samples for particulate
organic carbon and nitrogen (POC and PON) and C and N isotopic composition
were analyzed on an Elemental Analyzer Flash EA 1112 series (Thermo Fisher)
coupled to a continuous-flow isotope ratio mass spectrometer (Finnigan Delta
Plus XP, Thermo Fisher). Table 2 summarizes N2, and C fixation rate
measurements are given in the Supplement. Datasets were
deposited on PANGAEA.

Table 2CO2 and N2 fixation rates based on triplicate
measurements at stations 1 (17.9970∘ N, 88.9968∘ E), 4
(16.9828∘ N, 89.2063∘ E), and 5 (17.2075∘ N,
89.4282∘ E). N2 fixation was only measurable in two
individual samples but only in one out of three technical replicates.

DNA was extracted using an established protocol based on a phenol and chloroform
extraction (Giovannoni et al., 1996). The quality and
concentration of the purified DNA was checked spectrophotometrically and
using the Quant-iT PicoGreen dsDNA kit (Invitrogen, Carlsbad, USA).

A metagenome from the deep chlorophyll maximum (DCM; 84 m water depth) at
station 4 was sequenced with Illumina HiSeq using a 2 bp×125 bp read length on a
Nextera XT library at the Institute of Clinical Molecular Biology (IKMB) at Kiel
University, Germany. Sequencing resulted in 321 Mb. Sequences were analyzed
using the MetaPathways pipeline (Konwar et al., 2013), a modular
annotation and analysis pipeline for predicting diversity and metabolic
interaction from environmental sequences consisting of a quality control, an
open reading frame prediction and annotation, diversity analysis, and
environmental pathway reconstruction. Phylogenetic identification of operational taxonomic units (OTUs)
was derived via a comparison with the RefSeq and Greengenes databases
(DeSantis et al., 2006). After a quality check, 6454
sequences of ribosomal RNA were identified, 622 286 sequences (27.56 %) of
proteins with known functions were identified, and 1 628 841 sequences (72.15 %) were
predicted proteins with an unknown function.

nifH gene amplification was performed using a nested polymerase chain reaction (PCR) protocol
(Zehr et al., 1998). PCRs were performed using the GoTaq
kit (Promega, Fitchburg, USA), adding 1 additional microliter of bovine serum albumin (BSA; 20 mg mL−1; Fermentas, Waltham, USA). The Topo TA Cloning® kit
(Invitrogen, Carlsbad, USA) was used for cloning of PCR amplicons, according
to the manufacturer's protocol. Sanger sequencing (340 nifH sequences) was
performed by the Institute of Clinical Molecular Biology, Kiel, Germany.
Negative controls were performed using the PCR mixture as described without
template DNA; no amplification was detected. Samples from the particulate
fraction >3µm were consistently negative for nifH gene copies
and were thus not further investigated. Sequences were ClustalW aligned in
MEGA 7 (Kumar et al., 2016), and a maximum-likelihood tree was
constructed on a 321 base pair fragment. Reference sequences were obtained
using BlastX on the NCBI database. Sequences were submitted to GenBank
(submission ID 2245434). The metagenome was submitted to the NCBI's
sequence read archive (accession number SRR9696254).

Quantitative real-time PCRs for nifH were performed using the cluster-specific
TaqMan probe qPCRS as described in Löscher et al. (2014), with primers,
probes, environmental standards, and PCR conditions as presented in the text.
Samples were run in duplicates on a Bio-Rad qPCR machine (Bio-Rad, Hercules,
USA).

2.4 Box model exercise

We used a simplistic five-box representation of an upwelling system with a
deep- and intermediate-water iron source, with primary and export production
as well as respiration derived from the original models
(Canfield, 2006; Boyle et al., 2013). The model was
used to distinguish a N2 fixation state of the BoB and a non-N2
fixation state with primary production driven by recycled dissolved nitrogen
compounds. In contrast to the previous model versions, we applied a non-Redfield-based N2 fixation scenario. Ammonia concentrations were set to
zero in all boxes, in accordance with our direct measurements. Fe
concentrations were set to 0.1 µmol L−1 in the deep- and
intermediate-water boxes and 0.00044 µmol L−1 in the productive
zone (Grand et al., 2015a, b). Oxygen concentrations
were adjusted to our measurements, with 220, 0.02, and 50 µmol L−1
in the surface (corresponding to the upper 60 m of the water column), OMZ,
and deep water, respectively (Bristow et
al., 2017). Phosphate and nitrate concentrations were taken from our
measurements, with phosphate concentrations of 0, 2.7, and 2.5 µmol L−1 in the surface, OMZ, and deep boxes, respectively, and oxidized
nitrogen compounds (nitrate + nitrite) at a concentration of 0, 38, and 35 µmol L−1 in the surface, intermediate, and deep boxes,
respectively. Further information on the model stoichiometry is given in the
Supplement.

We explored the diversity, distribution, and activity of N2-fixing
microbes and carbon fixers in the OMZ of the northern BoB during the
northeast monsoon in January 2014. During the time of the cruise, low sea
surface temperatures (SSTs; surface waters refer to water depths shallower
than the mixed layer depth of 60 m) and low surface water salinity reaching
from the coasts of India, Bangladesh, and Myanmar southwards to approximately
16∘ N were present (Fig. 1a, b). At the coast, this low-salinity and low-SST plume co-occurred with increased chlorophyll concentrations
(Fig. 1c), thus suggesting a stimulation of primary production by waters
possibly of riverine origin (Fig. 1c). This is in line with earlier
suggestions of riverine-nutrient runoff promoting primary production close
to the shelf, where nutrients are consumed rapidly, thus preventing their
offshore transport (Kumar et al., 2004; Singh et al., 2012; Singh and
Ramesh, 2011; Krishna et al., 2016). Chlorophyll concentrations in the BoB
during the time of the cruise detected via satellite monitoring ranged
between 0.08 mg m−3 in open waters and 15 mg m−3 at the
northern coast and were consistent with previous in situ measurements during
low productivity periods in the BoB (Kumar et al., 2010).

The sampling stations were located offshore in the central BoB (Fig. 1),
where waters were strongly stratified with low sea surface salinity but
warmer SST compared to the coast and a steep oxycline reaching O2
concentrations close to anoxia at around 100 m water depth. No in situ
chlorophyll measurements are available from the cruise, but a fluorescence
sensor attached to the CTD showed a maximum of up to 0.8 mg m−3 between
32 and 90 m water depth (Fig. 2). Satellite-derived chlorophyll concentrations
in the coastal BoB were in the range from 0.08 to 0.35 mg m−3, slightly
higher than in a previous study of this region (0.06 mg m−3;
Kumar et al., 2002). Carbon fixation rates ranged between
286 and 1855 nmolCL-1d-1 at the depth of the DCM (Fig. 2, 84 m);
however, our rate measurements did not cover the water column above 60 m
water depth, where rates may have been higher. Consistent with previous
descriptions of primary producers at our study site (Loisel et al.,
2013) and with satellite imaging (Fig. S1 in the Supplement), we identified cyanobacteria
related to Synechococcus and Prochlorococcus as the most abundant primary producers in our
metagenome from the BoB DCM, accounting for 3.3 % of OTUs, while eukaryotic
phytoplankton accounted for only 0.3 % of OTUs (Table S1 in the Supplement).

Similar to chlorophyll, particulate organic carbon (Table S2; see also
Fig. S2 for a distribution of POC in the BoB) concentrations were low,
ranging between 4.96 and 7.84 µmol C L−1 in surface waters
and resulting in an average POC:chlorophyll ratio of 68:1 to 115:1 at the
depth of the DCM (Fig. 1). This ratio is comparable to POC:chlorophyll
ratios reported from cyanobacteria-dominated communities (74:1 to 126:1; e.g.,
Lorenzoni et al., 2015; Sathyendranath et al., 2009), but it is higher
compared to other OMZ regions (e.g., 50:1 in the eastern tropical South
Pacific; Chavez and Messié, 2009; Chavez et al., 1996).
Similarly, carbon fixation rates were 1–2 orders of magnitude lower compared
to the Arabian Sea, the tropical South Pacific, and the tropical Atlantic (e.g.,
Longhurst et al., 1995). While our POC concentrations from DCM
are 1 order of magnitude higher than the satellite-derived POC estimates
(Fig. S2) from surface waters, indicating that POC and primary production in
surface waters was not higher than in the DCM, it must be noted that our
measurements did not cover the entire mixed layer and are thus likely a
rather conservative minimum estimate.

Figure 3N:P ratio at station 1, 4, 5, and 6, with the Redfield ratio of N:P=16:1 indicated with a red line. The negative intercept of the trend line
indicates a deficit in dissolved inorganic nitrogen.

3.1N2 fixation in the upper water column and the oxycline

Based on the ratio of dissolved inorganic nitrogen (NO3-+NO2-) to phosphate (PO43-), which has a negative
intercept with the y axis (Fig. 3; Benitez-Nelson, 2000), primary
production in BoB waters appeared nitrogen limited during the cruise,
assuming Redfield stoichiometry. This nitrogen limitation would be expected
to create a niche for N2 fixation, but except for two samples for which
in both cases only one out of three technical replicates showed an isotope
enrichment, N2 fixation rates were below the detection limit (Table 2).
In this context, it is important to note that our rate measurements only
cover water depths between 60 and 280 m, thus excluding the upper part of
the euphotic zone. However, the absence of N2 fixation even in waters
shallower than 60 m is consistent with the observed δ15N
signatures (data available from 3 to 2300 m water depth;
Bristow et al., 2017) of both the
nitrate and the particulate organic nitrogen (PON) pool. δ15N
signatures were only slightly decreased in the top 100 m of the water column
to 5 ‰–8 ‰ (Fig. S3), thus not speaking for the presence of
active N2 fixation which would be expected to create substantially
lighter δ15N signatures of −2 ‰–2 ‰ (e.g.,
Dähnke and Thamdrup, 2013). Several clusters of N2-fixing microbes were, however, identified by screening for the key
functional marker gene nifH (Fig. 4). Only a few nifH sequences were associated with
cyanobacteria commonly abundant in ocean surface waters, even in the
euphotic zone at 10 m water depth. This pattern seems to be typical for OMZ
areas (Fernandez et al., 2011; Jayakumar et al., 2012; Löscher et al.,
2014) and for the eastern Indian Ocean (Wu et al., 2019), where
cyanobacterial nifH sequences are also rare. Similar to earlier studies, which
identified Trichodesmium in BoB surface waters (Bhaskar et al., 2007; Hegde, 2010; Wu et
al., 2019), we detected nifH copies related to Trichodesmium in our samples, both by
sequencing and by qPCR (Fig. 4; Table S3). These sequences clustered closely
to Trichodesmium and nifH previously recovered from the Arabian Sea (Jayakumar et al.,
2012; Mazard et al., 2004), where those N2 fixers were found in low
abundances, but possibly actively fixing N2, as indicated by nifH presence
in a cDNA library. No sequences related to the different groups of
unicellular cyanobacterial diazotrophs (UCYN-A, UCYN-B, or UCYN-C; Zehr et al.,
2001) were present in our nifH dataset. UCYN-A and UCYN-B have previously been
found in the Arabian Sea, but only at oligotrophic stations with warm water
temperatures >30∘C (Mazard et al.,
2004). While UCYN-A may occur at temperatures below 25 ∘C,
Trichodesmium and UCYN-B may be limited by the water temperatures at our sampling
stations, which were possibly too low, at around 25 ∘C.
Trichodesmium is usually abundant in high-iron-input regions such as the tropical Atlantic
Ocean (Martínez-Pérez et al., 2016). The absence
of Trichodesmium and other cyanobacterial N2 fixers may thus also result from an
insufficient iron source (Moore et al., 2013). Additionally, light
limitation due to severe atmospheric pollution (known as the South Asian
brown cloud) which lasts over the BoB from November to May (e.g.,
Ramanathan et al., 2007) may influence the distribution of
cyanobacteria in the BoB (Kumar et al., 2010). While earlier studies
also detected Chaetoceros (Bhaskar et al., 2007; Hegde, 2010; Wu et al., 2019), a
diatom known to live in association with diazotrophs, no diatom-associated
N2 fixers could be identified from our sequences. Thus our data do
not directly support previous suggestions of those specific diazotrophs
producing low δ15N nitrate signatures along with high opal
concentrations previously detected in sediment trap samples
(Gaye-Haake et al., 2005).

3.2N2 fixation in the OMZ

In the cruise area, we detected again the genetic potential for N2
fixation, but N2 fixation rates were below the detection limit and
δ15N signatures of nitrate and PON indicated nitrogen loss
instead of N2 fixation (Fig. S3). The community of N2 fixers in
the BoB consisted mostly of the non-phototrophic, proteobacterial
representatives of nifH – clusters I and III (Fig. 4), most of them related to
previously identified OMZ diazotrophs (Fernandez et al., 2011; Jayakumar
et al., 2012; Löscher et al., 2014).

Figure 5Venn diagram of nifH clusters present in Arabian Sea DNA libraries (AS)
and in cDNA libraries (AS cDNA) and clusters identified in the BoB,
O2-depleted basins of the Californian Bay (CB), the eastern tropical
North Pacific (ETNP), and the eastern tropical South Pacific (ETSP). Clusters
as depicted by triangles in Fig. S6 were collapsed based on a 98 %
identity. The black area shows the clusters present in all OMZs. Numbers
indicate the individual clusters in fields which would otherwise appear
unproportionally large.

A statistical comparison of BoB nifH sequences with OMZ diazotroph communities
from the Arabian Sea, the eastern tropical South Pacific (ETSP), eastern tropical
North Pacific (ETNP), and hypoxic basins in California Bay
revealed a strong similarity, suggesting that certain diazotrophs are
characteristic for OMZs (Fig. 5). Those typical OMZ clusters include
uncultured γ-, δ-, and ε-proteobacteria and clostridia. Only
one cluster was uniquely represented in the BoB and absent from the other
OMZ datasets, with only three individual sequences related to Azotobacter chroococum. Another
difference between the BoB and the other OMZ diazotroph communities was
the composition of Cluster IV nifH sequences, which are present but cluster in
different groups as compared to, for instance, the Arabian Sea Cluster IV
community. It is, however, unlikely that Cluster IV diazotrophs are
important for N2 fixation in the BoB or other OMZs because they were
never shown to be transcribed (Fernandez et al., 2011; Jayakumar et al.,
2012; Löscher et al., 2014), and Cluster IV nif is generally considered to
encode non-functional nif or paralogous sequences (Gaby and Buckley,
2014; Angel et al., 2018). In addition, the presence of Cluster IV nifH sequences
has previously been ascribed to PCR contamination (Zehr et
al., 2003). Thus, the importance of this cluster for N2 fixation in
OMZs is generally debatable, and the different composition of the Cluster IV
diazotroph community likely .does not explain the absence of N2 fixation in the BoB.

While diazotroph communities highly similar to the identified BoB
diazotrophs promote active N2 fixation in other OMZ waters, we have no
consistent indication for N2 fixation in the BoB (Table 2). One
explanation for the absence of N2 fixation could be the sensitivity of
the BoB OMZ diazotrophs to O2 as opposed to the relative O2 tolerance of cyanobacterial N2 fixers. We identified BoB diazotrophs
closely related to cultivated N2 fixers, including Vibrio diazotrophicus and Desulfonema limicola, which fix
N2 only under strictly anaerobic conditions (Urdaci et al.,
1988; Bertics et al., 2013; Gier et al., 2016). Further, communities of
diazotrophs from other OMZs highly similar to the BoB diazotroph community
were described to transcribe their nifH gene and to actively fix N2 only
under strictly anoxic or anoxic–sulfidic conditions (Löscher et al.,
2016, 2014; Jayakumar et al., 2012,
2017) and are unable to fix N2 in the presence of even minimal
concentrations of O2 (reviewed in Bombar et al.,
2016). N2 fixation in our samples (Table 2) may therefore be directly
inhibited by the detected traces of O2. Thus, our data suggest that
even only nanomolar O2 concentrations such as those present in the BoB may
prevent non-phototrophic N2 fixers from actively fixing N2, which
could ultimately limit the supply of new nitrogen to the BoB.

Figure 6Model of the response of the BoB OMZ to a weaker stratification
corresponding to increased upwelling in this model under a non-N2
fixation scenario with nitrate-driven production, photic-zone primary
production dependent on N2 fixation, and a scenario of N2 fixation in
the OMZ, which would result in build-up of a nitrogen stock and export to
the productive surface if stratification becomes weaker.

3.3 Role of Fe and mesoscale activities (eddies)

The high iron (Fe) requirement of N2-fixing microbes (60 times higher
compared to other marine organisms; Gruber and Galloway, 2008)
limits N2 fixation in large parts of the ocean (Moore et al., 2013).
However, eolian Fe fluxes to surface waters of the southern BoB were
estimated to be comparable to those detected underneath Saharan dust plumes
in the Atlantic (290±70µmolm-2yr-1;
Grand et al., 2015a). Indeed, dissolved Fe (dFe) accumulates
in the BoB OMZ, reaching comparably high concentrations of up to 1.5 nM
(Grand et al., 2015b; Chinni et al., 2019). In surface waters, dFe
concentrations were described to range from 0.4 nM in the area of the cruise
to up to 0.5 nM towards the north of the BoB, with increasing concentrations
coinciding with decreasing salinity north of 15∘ N (Grand et
al., 2015a, b; Chinni et al., 2019). While the reported Fe
concentrations do not indicate Fe limitation of N2 fixation in the
OMZ, surface primary production and N2 fixation may be limited by any
other micronutrient. Indication for such a limitation can be derived from
eddy-induced Ekman pumping; mesoscale dynamics and the summer monsoon
current have been shown to trigger plankton blooms with high productivity
(Jyothibabu et al., 2015; Vinayachandran and Mathew, 2003; Chen et al.,
2013; Fernandes et al., 2009), possibly induced by upwelling of certain
nutrients to surface waters. Besides locally increasing surface water
chlorophyll concentrations, erosion of the strong stratification and
subsequent nutrient input to surface waters result in a change of
phytoplankton size class (Prasanna Kumar et al., 2004).
While usually smaller phytoplankton dominate the primary producer pool (60 %–95 % of the total chlorophyll), the contribution of larger
phytoplankton has been observed to double in the regions influenced by the
summer monsoon current and in mesoscale eddies, which impacts the vertical
organic carbon flux in the BoB temporally and locally (Jyothibabu et al.,
2015; Prasanna Kumar et al., 2004; Huete-Ortega et al., 2010; Gomes et al.,
2016). The resulting increase in organic matter production, the modified
composition of organic matter (i.e., production fresh and labile particulate organic matter – POM), a
faster export, and subsequent respiration could promote anoxic OMZ conditions
in the BoB. This may subsequently allow for O2-sensitive processes to
take place, which may include N2 fixation and nitrogen loss processes
(Johnson et al., 2019), locally or regionally. Rapid changes in
dissolved O2 induced by increased surface productivity and organic
matter export were reported in the context of mesoscale water mass dynamics
in the BoB (Johnson et al., 2019) and also in other eddy
systems in the Atlantic, which showed rapid O2 exhaustion in otherwise
oxic waters (Fiedler et al., 2016; Löscher et al., 2015). Episodes of
increased biological productivity have also been reported from the BoB
during both the pre-southwest monsoon and northeast monsoon
(Kumar et al., 2004). Under those scenarios, large parts of the
BoB's surface waters exhibited a strong pCO2 undersaturation compared to
the atmosphere (∼ 350 µatm), resulting in an air–sea pCO2
gradient sometimes exceeding 100 µatm. This gradient is explainable
only by an increase in biological primary production fueled by temporal
external nutrient input (Kumar et al., 2004). As Singh et al. (2012) pointed out, these high-productivity episodes cannot be explained by
riverine or atmospheric deposition of nutrients alone, but upwelling or
N2 fixation would be required to sustain the nitrogen demand.

3.4 Feedbacks between N2 fixation and OMZ intensity

We used a simple model to test the conditions allowing for N2 fixation
in the surface waters and in the OMZ of the BoB and the interplay of
N2 fixation with primary production in response to changes in
stratification (i.e., upwelling). We further explored how far N2
fixation controls O2 concentrations in the BoB OMZ. We simulated
nitrate-driven primary production and primary
production dependent on N2 fixation, which is representative of N2 fixation in the photic zone
and governed by excess phosphorus and Fe availability as previously used in
Canfield (2006) and Boyle et al. (2013). In
addition, we simulated primary production that is dependent on
OMZ-associated N2 fixation, which, in contrast to the classical N2
fixation scenario, is independent of a Redfield-based nitrogen deficit, with
N2 fixation being active as long as phosphorus and Fe are available in
concentration >0 (Bombar et
al., 2016; Löscher et al., 2014). One weakness of this model simulation
is that it only includes Fe as potentially limiting nutrient for N2
fixation, which is, according to the available datasets (Grand et al.,
2015b; Chinni et al., 2019), not necessarily correct but may be valid as an
indicator for any other unrecognized micronutrient limitation. Consistent
with the previous deep-time models of Canfield (2006) and
Boyle et al. (2013), our model exercise revealed that
additional nitrogen supply by N2 fixation or other external nitrogen
sources would generally exhaust the remaining traces of O2 with
increasing upwelling (Fig. 6). According to our model, this would lead to
denitrification, which is in line with O2-manipulated experiments as
presented in Bristow et al. (2017) and consistent with the available isotope
records from the OMZ (Fig. S3). A weaker stratification (in the model
depicted as increased upwelling fluxes) would have the strongest effect on
oxygen exhaustion and the onset of denitrification if primary production is
dependent on N2 fixation in the photic zone, followed by OMZ-located
N2 fixation and lastly by nitrogen recycling. Given that OMZ regions are
sites of massive nitrogen loss characterized by a nitrogen deficit in the
water column (Deutsch et al., 2007), the similar
diazotroph community in the OMZ paired with an absence of N2 fixation
in the euphotic zone suggest that OMZ-associated N2 fixation is the
most likely scenario. Thus, nitrogen limited primary production in the BoB
and in OMZs in general would be susceptible to changes in stratification,
with increased upwelling of nutrient-rich waters causing O2 exhaustion.
Considering the potential O2 sensitivity of OMZ diazotrophs based on
the comparison with other OMZs, the interplay between O2
concentrations, stratification, and N2 fixation may act as a stabilizing
feedback on the BoB OMZ, preventing full O2 depletion.

One factor possibly disturbing a possible stabilizing feedback is the
external anthropogenic supply of nitrogen to the northern Indian Ocean. This
additional nitrogen source is projected to increase over the next decades
(Duce et al., 2008), potentially accelerating primary production in the
future ocean, including the BoB. An atmospheric input in the range of 1.1
(model based) to 1.6 Tg N yr−1 (observation based) has been reported,
which will likely increase in the future (Suntharalingam et
al., 2019). This additional nitrogen fertilization would cause the same
effect as N2 fixation in our model, thus exhausting the present
traces of O2 in the OMZ rapidly. Until an increased supply of
atmospheric or riverine nitrogen becomes significant, changes in water
column stratification, however, likely impose the strongest control on
N2 fixation and primary production and thus on respiration, nitrogen
loss processes, and ultimately on the O2 status of the OMZ in the BoB.

We detected a diazotrophic community similar to those from other OMZ
regions; however, we could not obtain consistent evidence for active N2 fixation in the BoB. Coming back to our original question of whether there is N2
fixation in the BoB, our data suggest that the answer is no. In other OMZs, N2
fixation has been observed to largely vary temporally and spatially, but
never reaching rates comparable to oligotrophic open ocean systems such as
the Pacific gyres. Episodes of N2 fixation, however, could be induced
by changes in water mass dynamics or riverine- or atmospheric-nutrient input.
Resulting increased N2 fixation and primary production would possibly
lead to O2 exhaustion in the BoB, which otherwise does not become fully
anoxic.

Previous observations describing the absence of nitrogen loss processes in
the BoB were explained by the remaining traces of O2
(Bristow et al., 2017) and possibly by a
nitrogen deficiency relative to carbon in the organic matter pool. While we
acknowledge that our dataset represents only a snapshot of the BoB's
biogeochemical setting, our observations may help in predicting the future
development of N2 fixation in the BoB and of the BoB OMZ with regard to
increased atmospheric dust deposition and ocean fertilization (Duce et
al., 2008), altered ocean circulation patterns (Yeh et al., 2009),
and deoxygenation of the ocean as a consequence of global warming
(Schmidtko et al., 2017; Stramma et al., 2008).

We thank the captain and crew of the ORV Sagar Kanya for their support during sampling.
We especially thank the Ministry of Earth Sciences (MoES), India, for
funding the research through the SIBER (INDIA) project GAP2425 and for
making ORV Sagar Kanya available for this work. We thank Julian Dekaezemacker and Laura Piepgras for sampling on board, for providing nitrogen and carbon fixation rates,
and for helpful comments on the dataset and Richard Boyle for providing the
backbone model. We thank Laura Bristow for helpful comments on an earlier
version of the paper, and we acknowledge Erik Laursen for technical
assistance, Cameron Callbeck and Gaute Lavik for sampling, and Alexander Treusch and Michael Forth
for providing access to subsamples for molecular analysis. We further thank
Gerd Krahmann for help with the analysis of fluorescence data from the CTD.
This study was supported by the Horizon 2020 program of the European Union (NITROX;
grant no. 704272 to Carolin R. Löscher) and the Max Planck Society. Further funding was
received from Villum Fonden (grant no. 16518; Donald E. Canfield) and the German Research
Foundation in the frameworks of the Cluster of Excellence “The Future Ocean”
and the Collaborative Research Center SFB754.

This research has been supported by the European Commission, Horizon 2020 (NITROX; grant no. 704272); the Villum Fonden (grant no. 16518); the Collaborative Research Center (grant no. SFB754); the German Research Foundation (DFG); and the Max Planck Society.

Christiansen, C. F. and Loescher, C. R.: Facets of diazotrophy in the OMZ
off Peru revisited: what we could not see from a single marker gene
approach, bioRxiv, 558072, https://doi.org/10.1101/558072, 2019.

Oxygen minimum zones (OMZs) are ocean areas severely depleted in oxygen as a result of physical, chemical, and biological processes. Biologically, organic material is produced in the sea surface and exported to deeper waters, where it respires. In the Bay of Bengal (BoB), an OMZ is present, but there are traces of oxygen left. Our study now suggests that this is because one key process, nitrogen fixation, is absent in the BoB, thus preventing primary production and consecutive respiration.

Oxygen minimum zones (OMZs) are ocean areas severely depleted in oxygen as a result of physical,...